ORIGINAL RESEARCH ARTICLES
DNA AND CELL BIOLOGY
Volume 33, Number 7, 2014
ª Mary Ann Liebert, Inc.
Pp. 399–407
DOI: 10.1089/dna.2013.2182
Mutations in the SPTLC1 Protein Cause Mitochondrial
Structural Abnormalities and Endoplasmic Reticulum
Stress in Lymphoblasts
Simon J. Myers,1,2,* Chandra S. Malladi,2,3,* Ryan A. Hyland,1,2 Tara Bautista,4 Ross Boadle,5,6
Phillip J. Robinson,7 and Garth A. Nicholson 8,9
Mutations in serine palmitoyltransferase long chain subunit 1 (SPTLC1) cause the typical length-dependent
axonal degeneration hereditary sensory neuropathy type 1 (HSN1). Transmission electron microscopy studies
on SPTLC1 mutant lymphoblasts derived from patients revealed specific structural abnormalities of mitochondria. Swollen mitochondria with abnormal cristae were clustered around the nucleus, with some mitochondria being wrapped in rough endoplasmic reticulum (ER) membranes. Total mitochondrial counts revealed
a significant change in mitochondrial numbers between healthy and diseased lymphocytes but did not reveal any
change in length to width ratios nor were there any changes to cellular function. However, there was a notable
change in ER homeostasis, as assessed using key ER stress markers, BiP and ERO1-La, displaying reduced
protein expression. The observations suggest that SPTLC1 mutations cause mitochondrial abnormalities and ER
stress in HSN1 cells.
Introduction
H
ereditary sensory neuropathy type 1 (HSN1) is
the most common dominantly inherited form of hereditary sensory neuropathy in man. The disorder has an onset in
late childhood, teenage, or early adult years and produces
sensory loss, which may go unnoticed. Loss of protective
sensation can lead to painless skin injuries and ulcerations
progressing to osteomyelitis and limb amputations. Degeneration of distal motor axons occurs later in life with distal
and later proximal limb weakness. Pathology shows early
sensory and motor distal axonal degeneration and later loss of
dorsal root ganglia and spinal motor neurons (Denny-Brown,
1952).
HSN1 mutations in the serine palmitoyltransferase long
chain subunit 1 (SPTLC1), cysteine (133) to tyrosine and
valine (144) to aspartic acid mutations (Dawkins et al., 2001),
may produce structural changes that can cause disease via
abnormal protein interactions. Whether these SPTLC1 mutations produce the HSN phenotype through loss of enzyme
activity is unclear. Dedov et al. (2004) and others (Bejaoui
et al., 2002) have demonstrated a loss of activity with the
cysteine (133) to tryptophan variant of human SPTLC1, as the
mutation is close to the active site of the enzyme (Dedov
et al., 2004). However, there are no changes in SPT products
in mutant SPT lymphoblasts (Dedov et al., 2004). The newly
described glycine (387) to alanine variant of human SPTLC1
(Verhoeven et al., 2004) is distant from the active site and is
unlikely to affect enzyme activity. Recent studies suggest that
these mutations cause a deleterious gain of function with the
production of toxic sphingolipids (Hornemann et al., 2009).
Patients suffering from HSN1 have an abundance of these
toxic sphingolipids in their blood stream, reflecting the observations of cell culture work.
Mitochondria play an essential role in peripheral nerves
and are implicated in some hereditary neuropathies (Niemann et al., 2005; Pedrola et al., 2005; Berger et al., 2006).
Mitochondria are transported down peripheral nerves and
cluster under areas of high-energy use such as internodes.
Mutations in mitofusin 2 produce the most common form of
1
Neuro-Cell Biology Laboratory, School of Science & Health, University of Western Sydney, Campbelltown, NSW, Australia.
Molecular Medicine Research Group, University of Western Sydney, Campbelltown, NSW, Australia.
Molecular Physiology Unit, School of Medicine, University of Western Sydney, Campbelltown, NSW, Australia.
4
Florey Neuroscience Institute, University of Melbourne, Victoria, Australia.
5
Electron Microscope Laboratory, Institute of Clinical Pathology and Medical Research, Westmead, NSW, Australia.
6
Westmead Millenium Institute, Westmead, NSW, Australia.
7
Cell Signalling Unit, Children’s Medical Research Institute, University of Sydney, Westmead, NSW, Australia.
8
Northcott Neuroscience Laboratory, ANZAC Research Institute, University of Sydney, Concord, NSW, Australia.
9
Molecular Medicine Laboratory, Concord Hospital, University of Sydney, Concord, NSW, Australia.
*These authors contributed equally to this work.
2
3
399
400
MYERS ET AL.
axonal Charcot-Marie-Tooth neuropathy (CMT type 2), and
CMT VI with optic atrophy. Mutations in mitofusin 2 disrupt
mitochondrial fusion (Legros et al., 2002) and mitochondrial axonal transport (Baloh et al., 2007). This is a lengthdependent axonal neuropathy where the longest axons are
affected first, commencing in the distal lower limbs and then
in the distal upper limbs. Since mitochondrial transport is
essential for distal axonal function, mitochondrial abnormalities have been proposed as the cause of a number of lengthdependent or ‘‘dying back’’ hereditary neuropathies.
Since structural abnormalities have been found in neuropathies, we examined mitochondrial morphology, mitochondrial number, mitochondrial length-to-width ratio, polarity of
mitochondrial membrane, and total cellular ATP levels in
lymphoblasts expressing the HSN1 mutation in SPTLC1. We
also monitored the expression of common ER stress markers
and measured receptor-mediated endocytosis. We propose that
cells expressing the SPTLC1 mutations have mitochondrial
structure abnormalities and a disrupted ‘‘ER homeostasis.’’
Methods
Cell culture
Control and patient lymphoblasts (harboring C133W or
V144D SPTLC1 mutations) were maintained in 5% CO2 and
at 37!C in RPMI-1640 (Gibco BRL) plus 10% fetal calf
serum (Gibco BRL), penicillin (100 mg/mL)/streptomycin
(100 units/mL) (Sigma-Aldrich), and 2 mM L-glutamine
(Sigma-Aldrich).
Electron microscopy
Control and SPTLC1 mutant lymphoblasts were washed
thrice before fixation in a modified Karnovsky fixative and
washed in 0.1 M 3-(-4-morpholino) propane sulfonic acid
(MOPS) buffer (pH 7.4). After washing, the cells were
postfixed in buffered 2% osmium tetroxide and dehydrated
in an ethanol series before embedding in Spurr epoxy resin.
Once polymerized (70!C for 10 h), sections were cut at
70 nm, stained with uranyl acetate and Reynold’s lead citrate and the sections were examined in a Philips CM120
BioTWIN electron microscope at 100 kV.
Indirect immunofluorescence and confocal microscopy
Lymphoblasts (1 · 106 cells) were suspended in 1 mL of
warm phosphate-buffered saline (PBS). After centrifugation
at 500 g for 5 min, the cell suspension was resuspended in
4% paraformaldehyde containing 0.3% Triton X-100 and
incubated at 37!C for 30 min. The cells were then centrifuged and blocked in 1% bovine serum albumin solution at
37!C for 30 min. After washing, the cells were resuspended
in primary antibody, MTCO2 (Abcam, 1:50), and incubated
for 1 h at room temperature. The cells were washed and
resuspended in secondary antibody, anti-mouse Rhodamine
(Millipore; 1:200), and incubated for 1 h at room temperature. DAPI stain (1 mg/mL) was added to the cell suspension
and after 2 min, the cells were centrifuged and washed
twice. Aliquots (200 mL) were added to microtiter plate
wells containing coverslips coated in Histogrip. The coverslips were washed in warm PBS, left overnight to dry, and
mounted onto glass slides before confocal imaging using
Zeiss Zen 2009.
FIG. 1. Representative transmission electron micrographs
(TEM) of cells from two different cultures of control and
serine palmitoyltransferase long chain subunit 1 (SPTLC1)
mutant lymphoblasts. (A, B) Control lymphoblasts displaying
typical large nuclei and mitochondria. In SPTLC1 mutant
lymphoblasts expressing the C133W mutations; lipid droplets
(arrowheads) and vacuole structures (arrows) are observed as
well as peri-nuclear localization of the mitochondria (C, D), the
endoplasmic reticulum (arrows) is also seen to be tightly
wrapped around the mitochondria (E, F). In SPTLC1 mutant
lymphoblasts expressing the V144D mutations; multi-vesicular
bodies (arrowheads) are present as well as peri-nuclear localization of the mitochondria (G–J). Scale bars 2 mM.
Flow cytometry
Aliquots of each cell suspension (40 mL) from section 2.3
were analyzed using the MACSquant flow cytometer (Miltenyi Biotech) and analyzed with MACSquant software. Live
MITOCHONDRIAL MORPHOLOGICAL CHANGES DUE TO MUTATIONS IN SPTLC1
single cells were gated, and the mean of fluorescence was
obtained per 10,000 events.
Mitoprobe JC-1 assay (mitochondrial
membrane potential)
Lymphoblasts (1 · 106 cells) were suspended in 1 mL of
warm PBS. Carbonyl cyanide 3-chlorophenylhydrazone
(CCCP; membrane uncoupler) (50 mM, 1 mL) was added to
the control tube and incubated at 37!C for 5 min. JC-1
(2 mM) was added to all the cells, and the cells were incubated at 37!C, 5% CO2 for 30 min. Cells were washed
once in PBS (2 mL), and then pelleted by centrifugation.
PBS (500 mL) was added to each tube, and samples were
analyzed by flow cytometry (BD FACscan) with 488 nm
excitation, where the depolarized mitochondria fluoresce
green.
Annexin V apoptosis assay
As per the manufacturer’s instructions (BD Biosciences
Pharmingen) briefly, lymphoblasts (1 · 106 cells/mL) were
washed twice in cold PBS and resuspended in 1 · binding
buffer. About 1 · 105 cells (in 100 mL) were transferred to a
5 mL culture tube, where 5 mL of Annexin V-FITC and 5 mL
401
of Propidium Iodide was added. Cells were gently vortexed
and incubated at room temperature for 15 min in the dark.
Binding buffer (1 · , 400 mL) was added to each tube, and
samples were analyzed by flow cytometry (BD FACscan).
Cellular ATP assay
The total cellular ATP level of the lymphoblasts was
measured using the ATPlite luminescence ATP detection
assay system kit (Perkin Elmer) according to the manufacturer’s instructions. Plates were dark adapted for 10 min, and
luminescence was measured using the luminometer (Victor
3 Multi Label Reader; Perkin Elmer).
Clathrin-mediated endocytosis assay
Lymphoblasts from healthy donors and from patients affected with HSN1 were cultured in RPMI that was supplemented with 10% FBS, 2% glutamine for 4 days. Clathrinmediated endocytosis (CME) was performed after 24 h serum
starvation in RPMI medium. Briefly, cells were incubated in a
5% CO2 incubator with Alexa Fluor 594 conjugated transferrin
(Molecular Probes) for 10 min and immediately washed with
acetic acid (0.2 M) with NaCl (0.5 M) for 10 min on ice. After
acid wash, cells were fixed in 4% r-formaldehyde for 10 min
FIG. 2. Representative TEM
displaying mutation specific ultrastructural changes from two different cultures of SPTLC1 mutant
lymphoblasts. Control lymphoblasts at 31,500 magnification show
a continuous outer membrane with
no breakages and are of normal size
(A, B). In SPTLC1 mutant lymphoblasts expressing the C133W
mutations; display a discontinuous
outer membrane and sections
where there is complete breakage
of the membrane (arrowheads) (C,
D). Arrows denote a curved membrane, which appears to bridge the
breakage. In SPTLC1 mutant lymphoblasts expressing the V144D
mutations; display electron dense
and grossly swollen cristae along
with discontinuous outer membranes (arrowheads) (E, F).
402
at room temperature. Cells were washed with cold PBS, and
the nucleus was stained with DAPI. Both control and patient
cells were spun on microscope slides (5 · 75 · 1 mm MenzelGlasser Super Frost plus; CE Invitro Diagnostics) using
Shandon Cytospin 4 (Thermo) at 800 rpm for 2 min. Transferrin uptake was measured in lymphoblasts and COS-7 cells
using MetaMorph Offline ver.6.2r1 software.
Western blots
Protein extracts (20 mg) from each lymphoblast sample
were electrophoresed on a 12.5% sodium dodecyl sulfate
polyacrylamide gel electrophoresis gels, transferred to polyvinylidene difluoride (0.2 mm) membranes, and probed using
the following antibodies: mouse anti-MTCO2 (Abcam;
1:2000), mouse anti-Actin (Millipore; 1:2,000,000), and
rabbit anti-BiP (Cell Signaling; 1:1000). Western blots were
developed using chemiluminescence and imaged using the
AGFA CP1000 X-ray processor (AGFA) followed by
analysis with Image-J by densitometry.
FIG. 3. Total mitochondrial
numbers and mitochondrial length/
width ratios. (A) % mitochondrial
numbers per cell was determined
by TEM (average of 100 cells) and
flow cytometry (MTCO2 stained
cells, mean of fluorescence peak
from 10,000 events). (B) Lengthto-width ratios of the mitochondria
counted in (A) (n = 100, – SD).
(C) Immunofluorescence of cells
stained with MTCO2 (Red),
and Dapi (Blue).
MYERS ET AL.
Results
Structural changes to mitochondrial morphology
in SPTLC1 mutant lymphoblasts
To observe the effect of the SPTLC1 mutations on cellular morphology, transmission electron microscopy (TEM)
was performed on control and SPTLC1 mutant lymphoblasts containing either the C133W or the V144D mutation.
Gross disruption of mitochondrial morphology was apparent
(Fig. 1). Control lymphoblasts display normal mitochondrial
morphology and normal structure (Fig. 1A, B). Lymphoblasts
carrying the C133W mutation have numerous lipid droplet
deposits (arrowheads), vacuolar structures (arrows) and there
appears to be reduced membrane ruffles in these cells (Fig. 1C,
D). At a higher magnification, the perinuclear-clustered mitochondria are ‘‘wrapped’’ in rough endoplasmic reticulum
(RER) in the cells expressing this C133W mutation (arrows)
(Fig. 1E, F). In cells containing the V144D mutation, the
membrane surface displays reduced ruffling as compared with
control cells. There are vacuolar inclusions and the appearance
MITOCHONDRIAL MORPHOLOGICAL CHANGES DUE TO MUTATIONS IN SPTLC1
403
of multi-vesicular bodies (MVBs; denoted by arrowheads,
Fig. 1G–J). The morphology-challenged mitochondria appear
to be peri-nuclear localized in the lymphoblasts carrying the
C133W mutation.
Figure 2A and B displays mitochondria in isolation from
control lymphoblasts that show a continuous outer membrane with no breakages, and they appear of normal size and
are not excessively electron dense. In cells expressing the
C133W SPTLC1 mutation, there is a discontinuous outer
mitochondrial membrane and areas of complete membrane
breakage (arrowheads) (Fig. 2C, D). In addition, arcs of
cristae membrane appear to form where full mitochondrial
membrane breakage occurs (arrows). In cells carrying the
V144D SPTLC1 mutation, disturbance to the continuity of
the outer mitochondrial membrane is also observed (arrowheads) (Fig. 2E, F). However, the major morphological
findings with this mutation are that the mitochondria are
extremely electron dense and have grossly enlarged cristae.
Total mitochondrial numbers in SPTLC1
mutant lymphoblasts
Due to the change in structure and localization of mitochondria in cells expressing the SPTLC1 mutations, total
mitochondrial number and length-to-width ratios were determined. Figure 3A displays total mitochondrial counts as
determined by TEM and flow cytometry, with each number
representing the % number of mitochondria per cell taken
from 100 cells or 10,000 cells respectively. The C133W
mutation in SPTLC1 increases mitochondrial number, but
this is not observed in the V144D mutation. Mitochondrial
length-to-width ratios revealed no difference between control and mutant SPTLC1 cells (Fig. 3B). Indirect immunofluorescence detection of the mitochondria-specific marker
MTCO2 (red) displayed no change to the cellular localization of the mitochondria between control and mutant
SPTLC1 cells (Fig. 3C).
Mitochondrial membrane potential in nonapoptotic
SPTLC1 mutant lymphoblasts
In order to determine whether the morphological changes
to the inner and outer mitochondrial membranes affect mitochondrial membrane potential a JC-1 assay was performed.
Figure 4A shows a graphical representation of polarized
versus nonpolarized mitochondria measured as a percentage
of cellular events from compiled flow cytometry scans. These
data show normal mitochondrial membrane potential for
control lymphoblasts (open bars) with drug-induced (CCCP)
depolarization of mitochondrial membranes in control cells
(negative control) displaying complete mitochondrial membrane uncoupling. However, in the overall pooled data from
the SPTLC1 mutant lymphoblasts, there was a small but not a
significant increase in membrane depolarization (shaded
bars). To determine whether this slight increase in mitochondrial membrane depolarization affected the overall ATP
content in the patient cells, total ATP levels were assayed.
There was no significant difference between control (open
bars) and patient (hatched bars) samples (Fig. 4B). To confirm that the cells were not actively undergoing apoptosis, the
Annexin V assay was performed. A histogram of control
(open bars) and patient (solid bars) lymphoblast cells shows
that the cells were not apoptotic (Fig. 4C).
FIG. 4. Intracellular energy levels and apoptosis profile in
SPTLC1 mutant lymphoblasts. (A) Mitochondrial membrane
potential was measured in SPTLC1 mutant lymphoblasts using the Mitoprobe JC-1 assay kit for flow cytometry. Data are
represented as a percentage of polarized mitochondria (open
bars) to depolarized mitochondria (solid bars) in a pooled
population (n = 3) and in individual patients (n = 1 but performed in triplicate). For each sample n = 3, – SD. (B) ATP
levels were assayed in control and SPTLC1 mutant lymphoblasts using the ATPlite luminescence ATP detection kit
(Perkin-Elmer). Data are represented as a fold change in ATP
production where the control cells are in open bars (n = 3) and
the pooled patient cells are hatched bars (n = 3). For each
sample n = 3, – SD. The individual patient cells are displayed
in solid bars (n = 1 but performed in triplicate). (C) The results
of an Annexin V assay where the data are shown as a percentage of cellular events for a pooled population of control
(open bars) (n = 3) and patient (solid bars) (n = 3) samples.
Live cell, early apoptotic, late apoptotic, and necrotic cell
populations are exhibited. For each sample n = 3, – SD.
404
Endocytosis in SPTLC1 mutant lymphoblasts
To determine whether the morphologically challenged
mitochondria affected other energy utilizing cellular events,
CME was assayed by the observed uptake of fluorescent labeled transferrin into the cell. The same amount of transferrin
(red) uptake was observed for control, C133W, and V144D
lymphoblasts (Fig. 5A–C). Single cell analysis shows no
variation in CME between control cells (Fig. 5D–G) and
C133W (Fig. 5H–K) and V144D (Fig. 5L–O) mutant cells.
Cells were counter-stained with the nuclear stain DAPI to
display the nucleus and show the localization of transferrinfilled endosomes. Quantitative analysis revealed no statistical
difference in CME between the three cell types (Fig. 5P).
Disruption to ER homeostasis in SPTLC1
mutant lymphoblasts
As observed in TEMs (Fig. 1), the dysfunctional mitochondria in the C133W mutant SPTLC1 patient cells are
wrapped in RER. Since the SPTLC1 protein is located in the
ER, we determined whether there is a changed expression of
ER stress marker proteins associated with these mutants
(Fig. 6). MTCO2, a marker for mitochondria, is unchanged
in these mutant cells (Fig. 6A, B). Interestingly, there was a
FIG. 5. Endocytosis in
SPTLC1 mutant lymphoblasts. Hereditary sensory
neuropathy type 1 (HSN1)
lymphoblasts have normal
receptor-mediated endocytosis (RME) function. (A–C)
Full-field views of control
cells, C133W, and V144D
patient lymphoblasts (respectively) where the nucleus
has been stained with DAPI
(blue), and CME was measured by the uptake of
fluorescent-labeled transferrin (red). (D–G) Single cell
images of control lymphoblasts; (H–K) single cell
images of C133W patient
lymphoblasts and (L–O) single cell images of V144D
patient lymphoblasts, all
showing normal uptake of
transferrin (red) and the
nucleus is stain with DAPI
(blue). (P) A graphical representation of transferrin
uptake measured as a percentage intensity of control,
C133W patient, and V144D
patient lymphoblasts (n = 400
cells per group).
MYERS ET AL.
significant increase in actin protein expression levels in
mutant SPTLC1 cells (Fig. 6A, C). In cells expressing the
C133W and V144D mutation in SPTLC1, BiP expression
levels significantly decreased and were almost undetectable
in V144D cells (Fig. 6A, D). A significant decrease was also
seen in ERO1-La, which is responsible for maintaining the
reduction potential of the ER (Fig. 6A, E).
Discussion
Our data indicate that the SPTLC1 mutations disrupt
mitochondrial morphology in lymphoblasts, by which the
ER wraps tightly around the mitochondria and ER stress
appears to be induced. Minimal loss of mitochondrial charge
is associated with the dysfunctional mitochondria. The
mechanism by which this process is occurring is unknown.
However, the apparent increase in stress due to SPTLC1
mutations may also make use of the mitochondria-ER
membrane juxtaposition (Martins de brito and Scorrano,
2010), leading to a collaboration between the mitochondria
and the ER for lipid exchange proteins to be transferred
across the organelles via the mitochondrial-associated
membrane (MAM) space and to dysregulate calcium efflux
to the mitochondria (Fig. 7).
MITOCHONDRIAL MORPHOLOGICAL CHANGES DUE TO MUTATIONS IN SPTLC1
405
FIG. 6. Protein expression
levels of several markers in
SPTLC1 mutant lymphoblasts. (A) Western blot
results of MTCO2: a mitochondrial marker, actin: key
component of the cytoskeleton, BiP: the key regulator in
protein folding and endoplasmic reticulum (ER) stress
and ERO1-La; responsible
for maintaining the redox
balance within the ER. (B)
The raw luminescence data
for MTCO2 (n = 3, – SD).
(C–E) The normalized data
for actin, BiP, and ERO1-La,
respectively (n = 3, – SD).
*p > 0.05, **p > 0.01.
The link between SPTLC1 mutations and the observed
mitochondrial changes is yet to be explored. The apposition
of the ER to mitochondria suggests a direct link (Martins de
brito and Scorrano, 2010). In general, decreasing the space
between the ER and mitochondria will cause an increase in
the concentration of Ca2 + within the MAM space, thus elevating mitochondrial Ca2 + levels. If one considers the dual
function of ER chaperone immunoglobulin heavy-chain
binding protein (BiP), first as a molecular chaperone and
inhibitor or the ER stress response, and second as an inhibitor of Ca2 + flux through the ER to the mitochondria
through Sigma-1 receptor and inositol trisphosphate receptor, then it is likely that small changes in BiP expression
levels could lead to large detrimental effects in mitochondrial form, as observed in the C133W SPTLC1 mutant, and
function. The observation in the V144D SPTLC1 mutant
that BiP expression is down-regulated offers another
mechanism of action, suggesting that this mutation may
negatively regulate the molecular pathway associated with
BiP. This possibly provides a functional link between ER
homeostasis and mitochondrial structure and function.
This is the first time that such a study has been performed
on HSN1 cells which show these ultrastructural changes to
mitochondria. In a yet-to-be-published study from our laboratory, other CMT types caused by mutations in mitofusin
2 and in dynamin 2 appear to present with different ultrastructural changes than those observed in these HSN1 cells
caused by the SPTLC1 mutations. In a study conducted by
Parra et al., (2008), it was shown that ceramide induces a
loss of mitochondrial connectivity and mitochondrial fragmentation, ultimately leading to apoptosis; this finding was
in direct contrast to our studies that exhibit no apoptosis.
Functional mitochondria are usually transported along
microtubules, with depolarized mitochondria returning to
the perinuclear region of the cell, suggesting that partially
depolarized mitochondria may not be transported distally
(Baloh et al., 2007). Therefore, a minimal loss in mitochondrial charge may be sufficient to disrupt mitochondrial
406
MYERS ET AL.
FIG. 7. A schematic representation of findings observed in this study showing
the liaison between the mitochondria, ER, calcium, and
SPTLC1.
transport, especially if extrapolated to the long distances
involved in a sensory neuron. Failure of the damaged mitochondria to reach the periphery may lead to changes in the
distal axon, which may cause axonal degeneration. The
precise degeneration mechanism is unknown but may make
use of normal developmental processes, such as retraction or
autophagy/mitophagy. These mitochondrial changes do not
appear to affect cellular apoptosis and obviously do not
affect CME in these cells. Likewise, they do not support an
active mitophagy process, as mitochondrial membrane potential would have been affected with depolarization of the
mitochondria (Min Jin and Youle, 2012). Ultrastructural
analysis should have displayed multi-lamellal (like) bodies
where the individual dysfunctional mitochondria have been
continuously wrapped in ER membrane or have engulfed the
ER membrane (Min Jin and Youle, 2012).
It is possible that the mutant SPTLC1 protein interferes
with the delivery of essential proteins, lipid, or Ca2 + from
the ER to the mitochondria (Bereiter and Voth, 1994; Wang
et al., 2007). The significant decrease in BiP and ERO1-La
protein expression in SPTLC1 mutants suggests that ER
stress may play a role in this disease. Since BiP is the key
regulator of protein folding and cellular responses to ER
stress, we suggest that the mutants have an affected ER
homeostasis which could lead to an abnormal interaction
between the ER and mitochondria. Whether this is related to
Ca2 + flux through the cell is unknown.
Both the ER and mitochondria bind to microtubules and
actin filaments, using the cytoskeleton as scaffolding for
shape and localization. The change in actin protein expression may help explain the close proximity of the ER to the
mitochondria through a change in the stabilization points
between the two organelles (Sturmer et al., 1995). The
change in actin is also important, as it may explain axonal
degeneration in sensory neurons carrying the SPTLC1 mutation through the inability of mitochondria to migrate to the
terminal end of the axon. An understanding of this process
may lead to therapeutic strategies, which may correct the
mitochondrial defect and, therefore, arrest the axonal degeneration found in this disease.
All of the hereditary neuropathies produce disability
through the common feature of axonal degeneration (Sahenk, 1999). Even initial demyelinating forms of neuropathy
eventually result in axonal degeneration (Madrid et al.,
1977). This is thought to be the result of disturbed Schwann
cell and axonal interactions and possibly the spread of highconductance sodium channels (Custer et al., 2003). Although much is known about the cell biology of many of the
neuropathy genes, little is known about how they promote
axonal degeneration in neurons.
This study provides evidence that SPTLC1 mutations
cause mitochondrial abnormalities and ER stress in cells
derived from patients. In other studies investigating hereditary neuropathies, mitochondrial abnormalities have been
suggested to be the causal mechanism (Abu-Amero and
Bosley, 2006; Kwong et al., 2006; Misaka et al., 2006;
Baloh et al., 2007), adding to the discussion that there is
potentially a mitochondrial trafficking defect within these
cells. How these changes cause distal axonal degeneration is
poorly understood. Various mechanisms have been proposed; one suggestion is the process of ‘‘localized apoptosis’’ (Carson et al., 2005). Here for the first time, we show
that cells expressing the SPTLC1 mutations have mitochondrial structure abnormalities and disrupted ‘‘ER homeostasis,’’ revealing a link between mitochondria, ER,
lipids, and calcium by which a common molecular mechanism appears close to being elucidated.
Acknowledgments
This work was supported in part by research grants from
USA-MDA (to G.N. and S.M.) and NH&MRC of Australia
(to P.J.R. and G.N.).
Disclosure Statement
No competing financial interests exist.
MITOCHONDRIAL MORPHOLOGICAL CHANGES DUE TO MUTATIONS IN SPTLC1
References
Abu-Amero, K.K., and Bosley, T.M. (2006). Mitochondrial
abnormalities in patients with LHON-like optic neuropathies.
Invest Ophthalmol Vis Sci 47, 4211–4220.
Baloh, R.H., Schmidt, R.E., Pestronk, A., and Milbrandt, J.
(2007). Altered axonal mitochondrial transport in the pathogenesis of Charcot-Marie-Tooth disease from mitofusin 2
mutations. J Neurosci 27, 422–430.
Bejaoui, K., Uchida, Y., Yasuda, S., Ho, M., Nishijima, M.,
Brown, R.H. Jr., Holleran, W.M., and Hanada, K. (2002).
Hereditary sensory neuropathy type 1 mutations confer dominant negative effects on serine palmitoyltransferase, critical for
sphingolipid synthesis. J Clin Invest 110, 1301–1308.
Bereiter-Hahn, J., and Voth, M. (1994). Dynamics of mitochondria in living cells: shape changes, dislocations, fusion and
fission of mitochondria. Microscopy Res Tech 27, 198–219.
Berger, P., Niemann, A., and Suter, U. (2006). Schwann cells
and the pathogenesis of inherited motor and sensory neuropathies (Charcot-Marie-Tooth disease). Glia 54, 243–257.
Carson, C., Saleh, M., Fung, F.W., Nicholson, D.W., and
Roskams, A.J. (2005). Axonal dynactin p150glued transports
caspase-8 to drive retrograde olfactory receptor neuron apoptosis. J Neurosci 25, 6092–6104.
Custer, A.W., Kazarinova-Noyes, K., Sakurai, T., Xu, X., Simon, W., Grumet, M., and Shrager, P. (2003). The role of the
ankyrin-binding protein NrCAM in node of Ranvier formation. J Neurosci 23, 10032–10039.
Dawkins, J.L., Hulme, D.J., Brahmbhatt, S.B., Auer-Grumbach,
M., and Nicholson, G.A. (2001). Mutations in SPTLC1, encoding serine palmitoyltransferase, long chain base subunit-1, cause
hereditary sensory neuropathy type I. Nat Gen 27, 309–312.
Dedov, V.N., Dedova, I.V., Merrill, A.H. Jnr., and Nicholson,
G.A. (2004). Activity of partially inhibited serine palmitoyltransferase is sufficient for normal sphingolipid metabolism
and viability of HSN1 patient cells. Biochim Biophys Acta
1688, 168–175.
Denny-Brown, D. (1952). The nature of muscular diseases. Can
Med Assoc J 67, 1–6.
Hornemann, T., Penno, A., Richard, S., Nicholson, G. van Dijk,
F., Rotthier, A., Timmerman, V., and von Eckardstein, A.
(2009). A systematic comparison of all mutations in hereditary
sensory neuropathy type I (HSAN I) reveals that the G387A
mutation is not disease associated. Neurogenetics 10, 135–143.
Kwong, J.Q., Beal, M.F., and Manfredi, G. (2006). The role of
mitochondria in inherited neurodegenerative diseases. J Neurochem 97, 1659–1675.
Legros, F., Lombes, A., Frachon, P., and Rojo, M. (2002).
Mitochondrial fusion in human cells is efficient, requires the
inner membrane potential, and is mediated by mitofusins.
Mol Biol Cell 13, 4343–4354.
Madrid, R., Bradley, W.G., and Davis, C.J. (1977). The peroneal
muscular atrophy syndrome. Clinical, genetic, electrophysiological and nerve biopsy studies. Part 2. Observations on pathological changes in sural nerve biopsies. J Neurol Sci 32, 91–122.
Martins de Brito, O., and Scorrano, L. (2010). An intimate
liaison: spatial organization of the endoplasmic reticulummitochondria relationship. EMBO J 29, 2715–2723.
Min Jin, S., and Youle, R. (2012). PINK1- and Parkin- mediated
mitophagy at a glance. J Cell Sci 125, 795–799.
Misaka, T., Murate, M., Fujimoto, K., and Kubo, Y. (2006). The
dynamin-related mouse mitochondrial GTPase OPA1 alters
407
the structure of the mitochondrial inner membrane when exogenously introduced into COS-7 cells. Neurosci Res 55,
123–133.
Niemann, A., Ruegg, M., La Padula, V., Schenone, A., and
Suter, U. (2005). Ganglioside-induced differentiation associated protein 1 is a regulator of the mitochondrial network:
new implications for Charcot-Marie-Tooth disease. J Cell
Biol 170, 1067–1078.
Parra, V., Eisner, V., Chiong, M., Criollo, A., Moraga, F.,
Garcia, A., Hartel, S., Jaimovich, E., Zorzano, A., Hidalgo,
C., and Lavandero, S. (2008). Changes in mitochondrial dynamics during ceramide-induced cardiomyocyte early apoptosis. Cardiovasc Res 77, 387–397.
Pedrola, L., Espert, A., Wu, X., Claramunt, R., Shy, M.E., and
Palau, F. (2005). GDAP1, the protein causing Charcot-MarieTooth disease type 4A, is expressed in neurons and is associated with mitochondria. Hum Mol Genet 14, 1087–1094.
Sahenk, Z. (1999). Abnormal Schwann cell-axon interactions in
CMT neuropathies. The effects of mutant Schwann cells on
the axonal cytoskeleton and regeneration-asociated myelination. Ann NY Acad Sci 883, 415–426.
Sturmer, K., Baumann, O., and Walz, B. (1995). Actin-dependent light-induced translocation of mitochondria and ER
cisternae in the photoreceptor cells of the locust Schistocerca
gregaria. J Cell Sci 108, 2273–2283.
Verhoeven, K., Coen, K., De Vriendt, E., Jacobs, A., Van
Gerwen, V., Smouts, I., Pou-Serradell, A., Martin, J.J.,
Timmerman, V., and De Jonghe, P. (2004). SPTLC1 mutation
in twin sisters with hereditary sensory neuropathy type I.
Neurology 62, 1001–1002.
Wang, L., Zhou, Y., Liu, B., Huang, S., Hou, F., and Ge, J.
(2007). Pro370Leu mutant myocilin disturbs the endoplasm
reticulum stress response and mitochondrial membrane
potential in human trabecular meshwork cells. Mol Vis 13,
618–625.
Address correspondence to:
Garth A. Nicholson, PhD
Northcott Neuroscience Laboratory
ANZAC Research Institute
University of Sydney
Hospital Road
Concord 2139
NSW
Australia
E-mail: [email protected]
Simon J. Myers, PhD
Neuro-Cell Biology Laboratory
Molecular Medicine Research Group
University of Western Sydney
Narellan Road
Campbelltown 2560
NSW
Australia
E-mail: [email protected]
Received for publication August 1, 2013; received in revised form February 21, 2014; accepted February 22, 2014.